Speciation Analysis of Butyl- and Phenyltin Compounds in Environmental Samples by GC

Separation and Atomic Spectrometric Detection

Dong Nguyen Van 2006

Umeå University Department of Chemistry Analytical Chemistry

Speciation Analysis of Butyl- and Phenyltin Compounds in Environmental Samples by GC

Separation and Atomic Spectrometric Detection

Dong Nguyen Van

AKADEMISK AVHANDLING

Som med vederbörligt tillstånd av Rektorsämbetet vid Umeå Universitet for erhållande av filosofie doktorsexamen framlägges till offentlig granskning vid Kemiska Institutionen, Sal KBC3A9, Fredag den 3 November 2006, Klockan. 10.00. Fakultetsopponent: Associate Professor Egon-Erwin Rosenberg, Vienna University of Technology, Institute for Chemical Technologies and Analytics, Department Instrumental Analytical Chemistry, Vienna, Austria

Title

Speciation Analysis of Butyl- and Phenyltin Compounds in Environmental Samples by GC Separation and Atomic Spectrometric Detection

Author

Nguyen Van Dong, Department of Chemistry, Analytical Chemistry, Umeå University,

SE-901 87 Sweden

Abstract The main goal of the work presented in this thesis is to improve the reliability of existing methods for speciation analysis of organotin compounds

Species-specific isotope dilution (SSID) calibration in combination with gas chromatography – inductively coupled plasma mass spectrometry was used to investigate the transformation of phenyltin species during sample preparation. Isotope-enriched phenyltin species were synthesized from corresponding isotope-enriched tin metals. SSID with a mixture of phenyltin species (PhTs) from one isotope was used to evaluate different extraction procedures for the determination of PhTs in fresh water sediment. Preparative liquid chromatography was used to produce single isotope-enriched phenyltin species making a multi-isotope spike (MI) SSID calibration possible. Different extraction procedures for the analysis of phenyltin species in biological samples were evaluated by applying MI-SSID. Degradation of TPhT and DPhT during sample extraction was observed and quantified. Accurate results were therefore obtained. A sample preparation procedure using mild extraction conditions with reasonable recoveries is described. The stability of organotin standards was investigated under different storage conditions. Mono- and diphenyltin were found to be redistributed and degraded during storage in methanol but were stabilized in sodium acetate/ acetic acid. A fast redistribution between monobutyl- and diphenyl tin has been observed and therefore it is therefore recommended that standards be derivatized as soon as possible after butyl- and phenyltin standards are mixed.

Included in the thesis is also an investigation of the analytical potential of using instrumentation based on atomic absorption spectrometry (AAS) for speciation analysis of organotin compounds. The method was based on gas chromatographic separation, atomization in a quartz tube and detection by line source (LS) AAS and for comparison, by state of the art continuum source (CS) AAS. Analytical performances of CSAAS system were found to be better compared to LSAAS.

Key words Organotin speciation analysis, species-specific isotope dilution, single isotope spike, multi-

isotope spike, redistribution, species transformation, preparative liquid chromatography, gas chromatography, line source atomic absorption spectrometry, continuum source atomic absorption spectrometry, inductively coupled plasma mass spectrometry

ISBN: 91-7264-187-8

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Speciation Analysis of Butyl- and Phenyltin Compounds in Environmental Samples by GC

Separation and Atomic Spectrometric Detection

Dong Nguyen Van

Department of Chemistry Analytical Chemistry

Umeå University

ii

Department of Chemistry Analytical Chemistry

Umeå University SE-901 87 Umeå, Sweden

Copyright © Dong Nguyen Van 2006 ISBN: 91-7264-187-8

Printed by VMC-KBC, Umeå, Sweden, 2006

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To Kim Oanh and my parents

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Title: Speciation Analysis of Butyl- and Phenyltin Compounds in Environmental Samples by GC Separation and Atomic Spectrometric Detection

Author: Dong Nguyen Van

Department of Analytical Chemistry, Umeå University S-901 87 Umeå, Sweden

Abstracts: The main goal of the work presented in this thesis is to improve the reliability of

existing methods for speciation analysis of organotin compounds Species-specific isotope dilution (SSID) calibration in combination with gas

chromatography – inductively coupled plasma mass spectrometry was used to investigate the transformation of phenyltin species during sample preparation. Isotope-enriched phenyltin species were synthesized from corresponding isotope-enriched tin metals. SSID with a mixture of phenyltin species (PhTs) from one isotope was used to evaluate different extraction procedures for the determination of PhTs in fresh water sediment. Preparative liquid chromatography was used to produce single isotope-enriched phenyltin species making a multi-isotope spike (MI) SSID calibration possible. Different extraction procedures for the analysis of phenyltin species in biological samples were evaluated by applying MI-SSID. Degradation of TPhT and DPhT during sample extraction was observed and quantified. Accurate results were therefore obtained. A sample preparation procedure using mild extraction conditions with reasonable recoveries is described.

The stability of organotin standards was investigated under different storage conditions. Mono- and diphenyltin were found to be redistributed and degraded during storage in methanol but were stabilized in sodium acetate/ acetic acid. A fast redistribution between monobutyl- and diphenyl tin has been observed and therefore it is therefore recommended that standards be derivatized as soon as possible after butyl- and phenyltin standards are mixed.

Included in the thesis is also an investigation of the analytical potential of using instrumentation based on atomic absorption spectrometry (AAS) for speciation analysis of organotin compounds. The method was based on gas chromatographic separation, atomization in a quartz tube and detection by line source (LS) AAS and for comparison, by state of the art continuum source (CS) AAS. Analytical performances of CSAAS system were found to be better compared to LSAAS.

Key words: Organotin speciation analysis, species-specific isotope dilution, single isotope

spike, multi-isotope spike, redistribution, species transformation, preparative liquid chromatography, gas chromatography, line source atomic absorption spectrometry, continuum source atomic absorption spectrometry, inductively coupled plasma mass spectrometry

ISBN: 91-7264-187-8

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List of papers

This thesis is based on the following papers and manuscript, which are referred to in the text by their Roman numerals.

I. Kumar S. J., Tesfalidet S., Snell J. P., Van D. N. and Frech W. A simple method for synthesis of organotin species to investigate extraction procedures in sediments by isotope dilution-gas chromatography-inductively coupled plasma mass spectrometry. Part 2. Phenyltin species J. Anal. At. Spectrom., 2004, 19, 368 – 372. http://dx.doi.org/10.1039/b313787b.

II. Van D. N., Muppala S. R. K., Frech W. and Tesfalidet S. Preparation, preservation and application of pure isotope enriched phenyltin species Anal. Bioanal. Chem., published online Aug. 26, 2006. http://dx.doi.org/10.1007/s00216-006-0695-8.

III. Van D. N., Bui T. X. T. and Tesfalidet S. Study on the degradation of phenyltin species during sample preparation of biological tissues using multi-isotope spike species-specific isotope dilution - GC-ICPMS Manuscript.

IV. Van D. N., Lindberg R. and Frech W. Redistribution reactions of butyl- and phenyltin species during storage in methanol J . Anal. At. Spectrom., 2005, 20, 266–272. http://dx.doi.org/10.1039/b416570e.

V. Van D. N., Radziuk B. and Frech W. A comparison between continuum- and line source AAS for speciation analysis of butyl- and phenyltin compounds J. Anal. At. Spectrom., 2006, 21, 708–711. http://dx.doi.org/10.1039/b518270k.

Papers I, IV and V are reprinted from Journal of Analytical Atomic Spectrometry with permission of The Royal Society of Chemistry (copyright 2004, 2005 and 2006, respectively. Paper II is reprinted from Analytical and Bioanalytical Chemistry with kind permission of Springer Science and Business Media, (copyright 2006).

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Abbreviations AcOH Acetic acid BT(s) Butyltin compound(s) Bu Butyl Cy Cyclohexyl DBT Dibutyltin DPhT Diphenyltin MBT Monobutyltin Me Methyl MPhT Monophenyltin OTC(s) Organotin compound(s) Ph Phenyl PhT(s) Phenyltin compounds(s) Pr Propyl TBT Tributyltin TCyT Tricyclohexyltin TET Triethyltin TMAH Tetramethylammoniumhydroxide TPhT Triphenyltin AAS Atomic Absorption Spectrometry AED Atomic Emission Detector AES Atomic Emission Spectrometer APCI Atmospheric Pressure Chemical Ionisation CCD Charge Couple Device CH-GFAAS Continuously Heated - Graphite Furnace AAS CS Continuum Source EI Electron Impact ESI Electrospray Ionisation ESR Electron Magnetic Resonance FWHM Full Width at Half Maximum GC Gas Chromatography GF Graphite Furnace ICP Inductively Coupled Plasma ID/SSID Isotope Dilution /Species Specific Isotope Dilution IE Ion exchange LC/HPLC Liquid Chromatography/High Performance Liquid Chromatography LOD Limit Of Detection LS Line source MS Mass spectrometry NP Normal phase P/FPD Pulsed/Flame Photometric Detector QF Quartz Furnace RF Radio Frequency RP Reversed phase SI/MI Single isotope spike/ Multi-isotope spike TOF Time Of Flight

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Tot- Total UV Ultraviolet XAFS X-ray Absorption Fine Structure Notations At Atomic Abundance Fi inter-conversion factor i m mixed or blend N number of mol R ratio of reference isotope signal to spiked isotope signal measured in the blend. s sample sp spike

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Table of contents

1 General considerations ...................................................................................................... 1

2 Organotin compounds........................................................................................................ 1

2.1 Chemical forms and usage ................................................................................................. 1 2.2 Organotin compounds in environmental systems ............................................................ 2 2.3 Pathways of organotin compounds in environment ........................................................ 4 2.3.1 Degradation ................................................................................................................ 4 2.3.2 Biomethylation ............................................................................................................ 4 2.4 Toxicity................................................................................................................................. 4 2.5 Legislation............................................................................................................................ 5

3 Techniques for organotin speciation analysis............................................................. 5

3.1 Direct techniques................................................................................................................. 5 3.2 Hyphenated techniques ...................................................................................................... 5 3.2.1 Liquid chromatography ............................................................................................... 6 3.2.2 Gas chromatography.................................................................................................... 7 3.2.3 Mass spectrometry and inductively coupled plasma mass spectrometry..................... 7 3.2.4 Atomic absorption spectrometry .................................................................................. 8

4 Analytical methodology for organotin speciation analysis .................................. 10

4.1 Gas chromatography-inductively coupled plasma mass spectrometry ....................... 10 4.2 Species-specific isotope dilution....................................................................................... 10 4.2.1 Principle of isotope dilution (ID) mass spectrometry ................................................ 10 4.2.2 Species-specific spiking strategies ............................................................................. 12 4.2.3 Preparation of the isotope-enriched phenyltin compounds. ...................................... 13 4.2.4 Species-specific isotope dilution to evaluate sample extraction procedures for PhTs16 4.3 Redistribution reactions of organotin compounds in standard solutions.................... 19 4.4 Gas chromatography atomic absorption spectrometry for OT speciation analysis ... 20 4.4.1 General...................................................................................................................... 20 4.4.2 Optimization of atomization conditions .................................................................... 21 4.4.3 Comparison between line- and continuum source AAS for organotin speciation .... 23

5 Conclusions ........................................................................................................................... 25

Appendix ................................................................................................................................... 26

Acknowledgements ................................................................................................................ 31

References ................................................................................................................................. 32

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1. General considerations It is well known that biological activities of an element and its effects on living organisms and

environment depend on its physiochemical forms. Analysts are therefore called upon to develop relevant and reliable analytical methods that make it possible to correlate the effects of a substance of interest in a sample with the composition of the sample matrix. To fulfill this goal, an analytical method must be able to provide sufficient information with known uncertainty about the identities and concentrations of the analytes in the sample. The aims of this thesis were to improve the reliability of current methods for speciation analysis of organotin compounds and to test possibilities to use a new technique based on continuum source (CS) atomic absorption spectrometry (AAS) for this type of application. According to the official definition of International Union of Pure and Applied Chemistry, speciation of an element is “the distribution of that element amongst defined chemical species in a system” and speciation analysis is “analytical activities of identifying and/or measuring the quantities of one or more individual chemical species in a sample”. The determination of total concentrations of the elements is usually insufficient for a satisfactory characterization of a sample. For example, the determination of total tin concentration in sediment contaminated with tributyltin (TBT) provides no meaningful data for risk assessment. In spite the fact that inorganic tin is rather harmless for the environment, TBT is known to have large adverse effects. An organometallic speciation analysis thus gives essential data for adequate risk assessment when combined with information about toxicity and transformation pathways of the species of interest in living organisms as well as in ecosystems.

The uncertainty of analytical results obtained by a speciation analysis method is governed by the least accurate and precise step in the whole analytical procedure. Sample preparation is likely the most critical step since it is prone to poor accuracy and precision. This holds true for the speciation analysis of organotin compounds, especially for phenyltin species, which are known to be unstable. Most of the studies in this thesis deal with the stability of organotin standards during storage and the degradation of phenyltin species during sample preparation, using conventional quartz furnace (QF) atomic absorption spectrometry and inductively coupled plasma (ICP) – mass spectrometry (MS) for detection. Some of studies deal with the investigation of the capability of organotin speciation analysis using a detector based on CSAAS. The performance of this detector was compared with the conventional line source (LS) AAS.

2. Organotin compounds

2.1 Chemical forms and usage Organotin compounds (OTCs) are implicitly referred to tetravalent organotin moieties which

are characterized by having a tin atom covalently bound to one or more organic substituents. These compounds are represented as RnSnX4-n, where R is any alkyl or aryl group and X is an anion such as halide, hydroxide, acetate etc. OTCs have been known since the middle of the eighteenth century. Their commercial applications were only widely recognized after 1940s when the plastics ndustry, especially the production of polyvinyl chloride (PVC), started to expand. Then the finding of the biocidal properties of trisubstituted OTCs in the late 1950s even broadened their application (see Table 1) [1]. The various commercial applications of OTCs resulted in a drastic increase in worldwide production of OTCs from less than 5000 tons in 1955 up to about 50000 tons in 1992 [2].

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Table 1: Main commercial application of OTCs

Application Function Compounds PVC stabilizers Stabilization against

decomposition by heat and light R2SnX2, RSnX3 R: Me, Bu, Oct

Antifouling paint Biocide R3SnX R: Bu, Ph

Agrochemicals Fungicide, insecticide, miticide, antifeedant

R3SnX R: Bu, Ph, Cy

Wood preservation Insecticide, fungicide Bu3SnX Glass treatment Precursor for SnO2

films on glass Me2SnX2, RSnX3 R: Me, Bu

Material protection Fungicide, algaecide, bactericide Bu3SnX Impregnation of textile Insecticide, antifeedant Ph3SnX Poultry farming Dewormer Bu2SnX2

About 70% of the total annual organotin production is used as additives for thermal and light

stabilization in the plastics industry as well as catalysts for polyurethane foams and silicones. The addition of alkyltin compounds, mostly mono- and dialkyltin, at concentrations of 5-20 g kg-

1 prevents the degradation of PVC under a prolonged exposure to heat and light. The use of TBT - based antifouling paints prevents the growth of aquatic organisms on vessel

hulls, which creates in the roughness of an increase in fuel consumption for example, a 10-µm increase in the rough average vessel hull may result in about 0.3-1% increase in fuel consumption [3].

In agriculture, tricyclohexyltin (TCyT) has been commonly used to control mites on fruits and triphenyltin (TPhT) has been used to control or prevent fungal diseases on a variety of crops [4]. On average, about 300 tons of TPhT are used annually in Netherlands for the treatment of potato crops and about 3 kg per hecta-acre per year of TPhT are used in the United States on pecan orchards.

2.2 Organotin compounds in environmental systems Owing to the widespread applications of OTCs, considerable amounts of these compounds

have entered various ecosystems, particularly the aquatic environment [5-7]. As Lewis acids, OTCs could receive lone pairs of electron to form neutral ion pairs (outer-sphere complex) or complexes (inner-sphere complexes).This changes the mobility and distribution of OTCs in eco-systems [8,9].

In general, the solubility of OTCs in water increases as the number and the length of the organic substitutes decrease. It also depends on the nature of the anion X, pH, ionic strength and temperature. Speciation of OTCs in aqueous solution is pH dependent. In aqueous solution, OTCs may exist as cations (below pH 4) or as neutral hydroxides (above pH 5) [10]. Enhanced levels of OTCs were found in waters adjacent to areas with high shipping activities like harbors (TBT), agricultural farms (TPhT) or in drinking water and municipal water (monobutyltin, dibutyltin) [11,12]. However, concentrations of TBT up to 20 ng L-1, which is ten times higher than the concentration believed to induce imposex in dogwhelk, were also detected in sea-surface micro layers and near-surface bulk water in open sea waters [13].

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In aquatic environment, tri-substituted OTCs have low solubility and low mobility and are easily adsorbed onto suspended particulate matter [14]. A large proportion of organotin contaminants are found to be associated with the clay fraction of particulate matter. This indicates that the adsorption and enrichment onto this fraction are important controlling mechanisms in regard to the distribution and fate of OTCs in the environment. Sorption of OTCs depends on pH, salinity and the mineralogical and chemical composition of the adsorption material. Processes of cationic exchange and complexation with negatively charged ligands such as carboxylate and phenolate are governing factors for the adsorption [9]. The presence of OTCs in sediments or suspended particulate matter makes OTCs available to filter- or sediment- feeding organisms. Hence long after the anthropogenic input of OTCs to the environment had been stopped, aquatic activities may still result in contamination of the aquatic system by re-suspension or remobilization of OTCs through dredging, swirling or desorption.

Since OTCs are lipophilic, they could extensively accumulate in fatty tissue. Tri-substituted OTCs are of most concern since they exhibit the highest toxicity and accumulation ability to organisms and animals [15,16]. The most important pathways by which an organism can take up OTCs, are bioconcentration and biomagnification. The former is defined as the uptake of OTCs from water or sediment phase via body surface and the latter is defined as the uptake of OTCs via the food chain. Accumulation is the result of both pathways and is often proportional to concentration of OTCs in the environment. The extent of bioaccumulation is also affected by biodegradation and/or excretion mechanisms of the respective organism.

OTCs were found in very high concentrations in aquatic living organisms, especially in invertebrates such as mollusk and barnacles, although their surrounding water contains only low ng L-1 levels of OTCs. TBT levels found in dogwhelk were 1000 times higher than the concentration in surrounding water [17]. High concentrations of BTs were found in trophic vertebrate predators or in birds, which are the higher trophic level of the food chain [18]. In birds, biomagnification factors for organotin were estimated to be in the range of 1.1-4.1 [19]. It is generally accepted that large bioaccumulation and biomagnification of organotin species induces the adverse effects to aquatic organisms and animals in the higher trophic level of the food chain even at very low concentrations of these pollutants in water.

The distribution of OTCs in environmental samples can give information about the contamination sources with respect to the history and type of OTCs as well as the physical/chemical conditions of the ecosystems. For example, the high levels of TBT in sediments at harbours were due to the release of antifouling paints from shipping activities in this area. However, high levels of tetrabutyltin (TeBT) and IOT found in Mulde River did not originate from antifouling paints but rather from untreated discharges from a nearby chemical plant [1]. In other findings, high concentrations of mixed butylmethyltin species in sediments of the river Elbe were not anthropogenically introduced into the aquatic environment but were presumably formed by methylation. Increase in the TBT gradient from the bottom to the top of a sediment core taken from the harbor of Arcachon indicates that TBT had persisted within the sedimentary column over a long time [1].

In biota, the higher ratios MBT/Tot-BTs and DBT/Tot-BTs in samples from industrial countries is correlated to high levels of MBT and DBT inputs, probably from the use of these agents as stabilizers in chlorinated polymers [20]. The predominance of TBT in deep-sea organisms may indicate a lower ability of organisms to decompose TBT as well as the inhibition of TBT degradation [21] by the prevailing environmental conditions such as low temperatures, lack of sunlight and phytoplankton.

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2.3 Pathways of organotin compounds in the environment

2.3.1 Degradation The degradation of OTCs in the environment can be described by successive losses of organic

groups as shown below. R4Sn → R3SnX → R2SnX2 → RSnX3 → SnX4

The breakdown of the organic groups occurs probably through various processes i. Ultraviolet radiation: photolysis by sunlight appears to be the fastest route of degradation

of OTCs in water. The mean bond dissociation energies for Sn-C bonds are in the range 190 to 220 kJ mol-1 while the energy for UV radiation at 290 nm is about 300 kJ mol-1. If OTCs absorb the sunlight, energetic photons provide sufficient energy to break Sn-C bonds.

ii. Biological cleavage: in the presence of bacteria or micro algae, TBT and TPhT biodegrade to less toxic di- and mono substituted species [22-24]. However, the biological activity of bacteria or micro algae is limited by many factors in their living conditions like temperatures, light or nutrients as well as the toxic concentrations of OTCs, which are biocide agents themselves.

iii. Chemical cleavage: the Sn-C bond can be attacked by both nucleophilic and electrophilic reagents such as mineral acid, carbocylic acid and alkalimetals.

Degradation of OTCs in the environment depends on their structures as well as various external conditions like matrices, temperatures, light etc. For example, in the presence of UV radiation, triphenyl- and tricyclohexyltin compounds are more rapidly degraded than TBT. Degradation of OTCs under UV radiation is particularly accelerated in the presence of Fe(III) or TiO2 as catalysts [25-27]. The half-life of TPhT depends on carbon content in soil [28]. Photolytic and microbial degradation of TPhT in soil were found to be slower compared to that in water [29].

2.3.2 Biomethylation Methyltin species found in the environment can be formed by biomethylation processes.

Methylcobalamin in the presence of Fe(III) or Co(III) can methylate inorganic or organic tin to monomethyltin or methybutyltin derivatives. Some bacteria, algae and seaweeds can also methylate Sn(II) to various forms of methyltin [30-32].

2.4 Toxicity Inorganic tin (IOT) compounds are generally considered nontoxic. However, the toxicity of an

OTC is known to depend on the nature and the number of organic groups that are bound to the tin atom. The toxic impact of a specific OTC on an organism is related to its exposed concentration and duration, bioavailability and sensitivity of the organism. In any RnSnX4-n series, tri-substituted OTCs are the most toxic. Among trialkyltin species, there are considerable variations in toxicity depending on the nature and side chain length of the alkyl groups. An increase in the n-alkyl chain length of a compound produces a sharp drop in biocidal activity. For example, triethyltin (TET) acetate is the most toxic compound of all organotins to mammals while trioctyltin (TOcT) compounds are essentially non-toxic to all organisms [33]. TBT and TPhT could cause adverse effects on aquatic organisms, even at low nanomolar aqueous concentrations [34]. Dibutyltin (DBT), which is highly neurotoxic, and monobutyltin (MBT) originating from the decomposition of TBT or from leaching of polyvinylchloride (PVC) are also of environmental concern [35].

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Tetra-substituted OTCs are nontoxic but delayed toxic activity is observed when they degrade to tri-substituted derivatives. Adverse effects of OTCs on humans such as decrease in thymocytes viability and lymphocyte concentrations in human blood have been observed [36]. In particular, triethyltin iodide, which was one of the ingredients in a drug, is believed to be responsible for over 100 lethal cases and 200 intoxications under the treatment of staphylococcal skin infections in France in 1954 [33].

The widespread applications of OTC have resulted in high risk of human exposure to these pollutants. Two pathways of OTC intake that have to be taken into consideration are ingestion of contaminated foodstuff (sea foods and drinking water) and indirect exposure to household items containing OTCs [37-40].

2.5 Legislation In 1982, laws regulating the use of organotin compounds were issued by France and later by

other countries; the use of TBT-based antifouling paints has been restricted for small ships (below 25m) and fish-farming equipment. The United Kingdom set an Environmental Quality Target Concentration for TBT in water at 2 ng L-1 based on the lethal concentrations for a few commercially important mollusks. In November 1999, the international maritime organization’s Marine Environment Protection Committee proposed a global prohibition on the application of organotin-based antifouling paints on ships by January 1, 2003 and a complete prohibition by January 1, 2008. Currently, the use of OTCs in European countries is subjected to the restrictions under commission directive 2002/62/EC, July 9, 2002.

Although phenyltin compound (PhTs) toxicity on various non-target organisms is well known, PhTs are still used in agriculture under certain restrictions. The World Health Organization (WHO) pronounced TPhT compounds as “safe agriculture chemicals” considering that their concentrations on treated plants decrease rapidly due to photo-degradation in the atmosphere besides being dissipated by wind or rain. Also, WHO argued that the residues of OTCs in foods, vegetables and fruits could partly or completely be removed by washing, peeling or cooking before consumption. However, it should be noted that recent studies have shown that cooking is not an effective way to eliminate OTCs from food [40,41].

3. Techniques for organotin speciation analysis

3.1 Direct techniques Direct techniques including Mössbauer spectrometry [42] and synchrotron-based X-ray

absorption fine structure spectrometry (XAFS) [43] have been used to provide information on electron structure, symmetry and the oxidation state of the organotin species and the local atomic arrangement around the tin atoms. This information is extremely useful for characterizing the distribution of OTCs in a sample giving rise to a better understanding of the pathway of OTCs in ecosystems. However, due to the lack of sensitivity, typically at the µg g-1 range, these techniques are not suitable for organotin speciation analysis in most environmental samples.

3.2 Hyphenated techniques Hyphenated techniques, also called indirect techniques, combine a chromatographic separation

and a detection system. The system combines the advantages of high separation efficiency of chromatography and high selectivity (element-specific or species-specific) and high detection power of a detector allowing speciation analysis at trace and ultra-trace levels.

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3.2.1 Liquid chromatography High performance liquid chromatography (HPLC) offers many advantages; it allows

separation of compounds with diverse polarity and volatility without the need of derivatisation. The risk of thermal decomposition of the analyte is small. There are several reviews on the application of HPLC for the speciation analysis of OTCs [44-46]. Separation of OTCs using ion exchange (IE), reversed phase (RP) or normal phase (NP) modes has been reported. Ion exchange chromatography is carried out on ionisable analytes using stationary phases that contain charge bearing functional groups and ionic-buffered mobile phases. Reverse phase separation mode uses a non-polar alkyl bonded silica-based stationary phase and a polar eluent. The normal phase mode uses polar bonded stationary phase and a relatively non-polar organic mobile phase. Various detectors for HPLC separation of OTCs using conventional or state-of-the-art spectrometers have been described. A summary of HPLC methods selected from the literature for the determination of OTCs is presented in Table 2. HPLC coupled to ultraviolet and fluorescence detectors for the determination of OTCs employing direct detection (for TPhT) or indirect detection has been described. For indirect detection, the on-line or off-line derivatisation of OTCs by a suitable fluorescent [47] or photometric reagent [48] was performed. A limitation of these methods is non-specific detection. Coupling HPLC to an element-specific detector, like flame or continuously heated (CH) graphite furnace atomic absorption and flame emission spectrometry, has been widely used for OTCs speciation analysis. The poor detection limit of these systems, due to low efficiency of nebulization and matrix interferences could be circumvented to some extent by using hydride generation, which, however, can only be applied to volatile hydride methyl- or butyltin compounds (BTs). HPLC coupled with for example ICPMS and atmospheric pressure chemical ionization (APCI)–MS have been extensively used in recent decades [49]. They show high sensitivity, large linear range, and isotopic selectivity.

Table 2: Summary of various phases HPLC methods for organotin speciation

Species; Sample matrix

Column Mobile phase Detector

LOD* (ng Sn)

TMT, TET, TPrT, TBT, TPhT; ground and estuarine water

Partisil SCX; (IE)

MeOH (70%), AcONa (0.01M), benzyltrimethylammonium chloride 0.002M

Indirect UV [44]

4 (TPhT) 100-820 ( others)

DBT, TBT Wood preservatives

Partisil SCX-10 (IE)

MeOH:water (60:40) NH4-citrate 0.66M MeOH:water (90:10), NH4OAc 0.66M

CH-GFAAS [50]

0.5 (TBT)

Sn(IV), TBT, TPhT, Water

Adsorbosphere SCX (IE)

MeOH 80%, Ammonium acetate 0.1M ICP-AES [44]

450-1500

DBT, TBT; PACS-1 Partisil SCX (IE) MeOH 60%, Diammonium citrate 0.18M ICPMS [44] 0.02-0.04MTs, BTs, DPhT, TPhT, sea water

TSK gel ODS-80TM (RP)

THF/water/AcOH: 54/38/8, tropolone 0.2% Long tube F-AAS [44]

5

DBT, TBT, DPhT, TPhT; Sea-waters and oyster tissues

Kromasil-100 C18 (RP)

Eluent A:MeOH/water/AcOH/TEA (57.5/2.5/40/0.05) + oxalic acid 30 mg L-1

Eluent B: MeOH/water/AcOH/TEA (84/1/15/0.03)

Indirect fluorometry [51-53]

0.02-0.5

BTs; standard TSK gel ODS-80TM (RP)

MeOH/water/AcOH: 80/14/6, tropolone 0.1%

ICPMS [54] 0.15-0.24

BTs, PhTs; PACS-2, habor sediment

TSK gel ODS-80TM (RP)

MeOH/water/AcOH/TEA: 72.5/21.5/6/0.1, 0.075% tropolone

ICPMS [55] 0.2-0.5

DBT, TBT, DPhT, TPhT, Sediment

Kromasil-100 (RP)

ACN/water/AcOH/TEA: 65/25/10/0.1 APCI-MS [56]

2.5-5

DBT,TBT, DPhT, TPhT, Sediment

Zorbax SB C18 (RP)

H2O-H2O/MeOH, TFA (1%), gradient APCI-MS [57]

0.02-0.07

DBT, TBT; Marine sediment

Nucleosil (NP Toluene 100%, tropolone 0.001% GF-AAS [58] 5 (TBT)

* Limit of detection is defined as three times the standard deviation of the noise

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As can be seen in Table 2, detection limits of OTCs for the various detection techniques are relatively poor. The reason for this is the inherently broad HPLC peaks and poor sample introduction efficiency to the detectors. Compared to the corresponding detection limits of OTCs for gas chromatography (GC), presented in Table 3, an improvement of 2-3 orders of magnitude is achieved for GC. The reason for this is that GC provides higher resolving power and complete sample transportation, thus improving sensitivities. For these reasons, hyphenated techniques using GC are generally preferred to HPLC for OTC speciation analysis.

3.2.2 Gas chromatography Gas chromatography (GC) has been extensively used as a separation technique for organotin

speciation analysis. Its high resolution allows the separation of many OTCs such as methyl-, butyl-, phenyl-, octyltin etc. in a sample. In addition, GC offers the ability to use one or more internal standards and surrogates, which permit the speciation of an analyte in every step of the analytical procedure to be traced. Since, separation on GC can only be applied for volatile compounds; derivatisation is required for most ionic, non-volatile OTCs. Thermally labile compounds cannot be analyzed by this technique. Hyphenated GC systems rely mainly on optical or mass spectrometric detectors. Atomic absorption spectrometry combined with quartz furnace (QFAAS) or atomic emission spectrometry (AES) and (pulsed) flame photometry ((P)/FPD) show moderate to high sensitivities for OTCs and they are rather inexpensive, user-friendly and commonly available in many laboratories. Therefore, these techniques are still useful. GC-MS is still very important since it is an indispensable tool for the identification of unknown species when investigating species transformation. With isotopic selectivity, the reliability of analytical results obtained by GC-MS/ICPMS can be improved because errors arising from incomplete extraction/derivatisation, instrumental drift, species transformation etc. can be addressed and corrected using species-specific isotope dilution methodology. The limits of detection for these detection techniques for organotin analysis are summarized in Table 3.

Table 3: Typical limit of detection for OTCs (pg as Sn) determined by GC coupled to different detectors

Detector QF-AAS FPD PFPD AES MS ICPMS LOD* (pg Sn) 17-37 [59] 4-10 [60] 0.07-0.38

[60] 2-7 [61]

0.1-1.0 [60] 0.2-1.0 [61] 0.05-0.08 [60]

* Limit of detection is defined as three times the standard deviation of the noise

3.2.3 Mass spectrometry and inductively coupled plasma mass spectrometry A mass spectrometric technique generally consists of an ion source and a mass analyzer. The

chemical information obtained from and the analytical performance of a mass spectrometric analysis depend on the ion source and the mass analyzer of the instrument. Molecular or elemental ions generated in the ion source are extracted, separated based on their mass to charge ratio (m/z) and counted. Various common mass analyzer designs involving quadrupole (Q)-, time of flight (TOF)-, magnetic sector field (SF)-, and ion-trap (IT)-MS and their combinations like Q-TOF-MS are available. Ion sources can be classified into two main groups: “soft” ion sources that provide low energy for ionization of molecular substances and “hard” ion sources that provide high energy to atomize and ionize the analytes with high efficiency. The soft ion sources such as electron impact (EI), chemical ionization (CI) or electrospray ionization (ESI) are useful for characterizing inorganic/organic compounds. The application of soft ion sources for quantitative analysis of organic compounds, especially of organometallic compounds, has been reported. One limitation of this approach is the relatively complex isotopic patterns

- 8 -

obtained from molecular species containing not only metallic element but also other elements like C, H, N. This complicates the calculation of the results when using species-specific isotope dilution (SSID). Hard ion sources like glow discharge and ICP can circumvent this problem by decomposing the organometallic compounds.

ICP is widely used for organometallic speciation analysis since it is compatible with continuous sample introduction and hence easy to couple to GC or HPLC. A schematic drawing of typical ICP-QMS is shown in Fig. 1. The ICP atomizer/ionizer contains a torch consisting of three concentric quartz tubes positioned centrally in a radio frequency (RF) coil, approximately 2-3 mm from the outer surface of the torch. The RF coil is connected to a RF generator operating at either 27 or 40 MHz and a power of typically 750 – 1600W to generate a strong magnetic field. Argon, which passes through the torch between the three concentric tubes at total flow rates typically 14-19 L min-1, is used to form a plasma discharge. The ICP discharge is sustained within the torch by continuous RF energy provided by the generator via the induction coil. Owing to the high ionization potential of argon (15.76 eV) and high RF power, plasma temperatures up to 10000 K can be obtained, which is sufficient to efficiently atomize and ionize most of the elements in the periodic table. Ions generated from the plasma are extracted by a reduced pressure through an ion source/mass spectrometer interface, which is maintained at a vacuum of 10-4 Pa using a roughing pump. The ions are then directed through ion optics placed between the skimmer cone and the mass analyzer where they are focused, accelerated and steered to the mass analyzer while particulates, neutral species and photon are removed. Quadrupole-based mass analyzer is at present the most commonly used (90% of all mass spectrometers). A quadruple used in ICP-QMS instrument operates at a frequency of 2-3 MHz and consists of four cylindrical stainless steel or molybdenum rods, which are typically 15-20 cm long and 1 cm in diameter. By applying variable direct current- and alternating current at a given setting of potentials on the quadrupole, only ions of a selected mass to charge (m/z) will pass through the space confined by the four rods with a stable flight path to reach the detector.

Figure 1: System overview of an Agilent ICPMS.1 GC, 2 torch, 3 induction coil, 4 sample cone, 5 skimmer cone

6 ion optics, 7 quadrupole, 8 detector, 9 rotary pump, 10 turbo pump. (Reprinted and adapted with permission from Agilent).

3.2.4 Atomic absorption spectrometry Conventional atomic absorption spectrometry nowadays can be considered an established

technique with no dramatic improvements to be expected in the future. The only real progress in the field of AAS is in the direction of high-resolution continuum source AAS, which could improve this technique. Atomic spectroscopic techniques are based on the measurement of

- 9 -

radiation at element-specific wavelengths, which is absorbed or emitted by free atoms of specific elements. Atomic absorption spectrometry has been used for the determination of metals and metalloids since 1952. An atomic absorption system generally comprises four main components; a primary light source, an atomizer, a mono or polychromator and a detector. The sample in either gas, liquid or solid form is introduced into an atomizer where it is thermally vaporized, atomized and, to a much lesser extent, ionized by flames or electrically heated cuvets. The free atom of analyte attenuates the specific radiation emitted from the light source. The non-absorbed radiation from the light source is directed through a mono- or polychromator so that only the selected monochromatic radiation with a narrow spectral bandwidth corresponding to the absorption line of the analyte will reach the detector where it is monitored. The absorption, which is proportional to the concentration of the free atom density in the atomizer, is calculated from the difference in radiation intensity in the presence and absence of atoms.

The instrumental features and operation conditions of conventional line source atomic absorption spectrometers have been described in several books [62-64].

3.2.4.1 Continuum source atomic absorption spectrometry The term “continuum source” describes a radiation source, which generates a spectrum of a

continuous spectral distribution over a broad wavelength range. The continuum source can be a high-pressure xenon short-arc lamp operating in the so-called “hot spot” mode. At a specific wavelength interval ∆λ, radiation emitted from the hot-spot that passes through the spectrometer is at least 10 times more intense compared to a corresponding emission lines from a hollow cathode lamp. The use of a double echelle monochromator (DEMON) provides enormous instrumental resolving power of 75000 or a spectral resolution of λ/∆λ=140000 [65], which corresponds to a full width at half maximum (FWHM) of the instrument profile of 2.7pm at 200 nm and 6.7 pm at 500 nm. The CSAAS employs a multi-channel solid-state array detector, typically a charge-coupled device (CCD), which offers the effective registration of spatially and temporally resolved intensity distributions, large dynamic range and high quantum efficiency in the UV region. A typical high-resolution CSAAS setup is illustrated in Fig. 2. CSAAS offers various advanced features compared to LSAAS;

i. The CCD array detector consists of several hundred pixels, which can be considered as independent detectors, simultaneously measuring radiation from the same light source. Only a few pixels are used to measure at the atomic absorption line profile, other pixels can be used to correct for fluctuations in the light source, to measure off the atomic absorption line profile.

ii. The wavelength-resolved detection of the atomic and background absorption gives more adequate information to correct for non-specific absorption. Background correction is performed based on truly simultaneous measurement of the background and analytical signals.

iii. The CSAAS provides relatively constant and highly intense emission (1 to 2 orders of magnitude compared to HCL) with very narrow spectral bandwidths in the whole spectrum of 190-900 nm range. This results in wavelength independent noise for all analytical lines with the possibility to select the analytical lines suitable to the analyte contents in a sample. Sample dilution is therefore not needed.

iv. CSAAS offers theoretically no limit on calibration range. In reality, the working range obtained with CSAAS is up to 5-6 orders of magnitude compared to 2-3 orders using magnitude of LSAAS. An improved detection limit by factors of 3-10 for CSAAS against LSAAS has been demonstrated [66].

- 10 -

Figure 2: HR-CSAAS setup with DEMON spectrometer. 1 Xenon short-arc lamp, 2 hollow cathode lamp (optional),

3 elliptical mirrors, 4 atomizers, 5 entrance slit, 6 parabolic mirrors, 7 prism, 8 folding mirrors and intermediate slit, 9 echelle grating, 10 CCD detector. Reprinted and adapted with permission from Ref.65.

4. Analytical methodology for organotin speciation analysis

4.1 Gas chromatography-inductively coupled plasma mass spectrometry The coupling of GC-ICPMS combines advantages of highly separating capability of GC,

excellent sensitivity and selectivity of ICPMS detector and 100% transportation efficiency of sample to the ionization source. The coupled system enables the determination of ultra trace levels of analytes in environmental samples with small needs for sample enrichment and clean-up. For organotin speciation analysis, the interface between GC and for example EI-MS or ICPMS should be kept at sufficiently high temperatures to prevent band broadening. For ICPMS, oxygen at a flow rate of several mL min-1 is added to the plasma to avoid the deposition of carbonaceous residues on the sampling cone of the MS. In contrast to GC-MS, the GC-ICPMS does not directly provide species-specific information and the identification of the species is based on the retention time.

4.2 Species-specific isotope dilution (SSID)

4.2.1 Principle of isotope dilution mass spectrometry Generally, isotope dilution methodology can be applied only for elements having at least two

stable isotopes. For each analyte species, a corresponding isotope-enriched analogue is used as internal standard for quantification. An accurately known amount of the isotope j-enriched analogue is added to the sample. The ratio of the two isotopes i and j, (i resulting from the analyte and j from the spike) in the mixture (blend) is measured. The same ratios for the individual spike and sample are also calculated based on their known isotopic compositions. The concentration of the analyte in the sample is calculated by comparing these three ratios according to equation 1.

)1(.

.. ,/,/

,/,/

sjimjim

mjimspji

s

sp

sp

ssps RRk

RkRmw

mAWAWCC

−

−=

- 11 -

Cs and Csp are concentrations (as Sn, w/w) of the analyte (unknown) in the sample and the spike (known); ms and msp are masses of the sample and the spike; w is dry weight correction factor. AWs and AWsp are average atomic masses of an analyte in the sample and in the spike;

isAt and i

spAt are the abundances (%) of reference isotope (i) in the sample and in the spike; j

sAt and jspAt are abundances (%) of enriched isotope (j) in the sample and in the spike; km is a

mass bias correction factor; mjiR ,/ is the ratio of reference isotope signal to enriched isotope

signal measured in the blend, e.g. ⎟⎟⎠

⎞⎜⎜⎝

⎛

Sn

Sn

j

i

SS

.

Assuming that equilibration of the spike and the analyte isotopes is achieved and that the isotope enriched analogue and incipient have the same physical and chemical properties, any change in concentrations of the analyte and the spike due to incomplete extraction, volatilization, etc, will affect both isotopes to the same extent. Total recovery of the analyte for an analytical procedure is not required because the measurement of the isotopic incipient/spike ratio cancels out all the changes.

When an isotope-enriched species (B), which is not spiked, is observed in the sample, it is likely that this species is formed as result of a transformation of another isotope-enriched species (A) in the spike i.e. species transformation from A to B has occurred. Quantification of species A and B allows characterization of species transformation and correction for changes in concentrations due to this process.

The isotope dilution methodology is less time consuming and can provide more reliable results than other calibration approaches such as external calibration, standard addition or internal standard calibration.

Isotope dilution techniques were first developed for elemental analysis during the 1950s [67]. This technique was extended into the field of analysis of organic compounds in the 1970s [68] and became a useful tool in analytical chemistry as a reference technique as well as for routine work e.g. residue analysis of dioxins [69]. At present, the ID methodology is also applied for speciation analysis of chromium, bromium, selenium, thallium, tin, iodine, platinum, lead and mercury [70]. Tin speciation analysis using species-specific isotope dilution has been focused on BTs and to some extent on PhTs. The approach has also been applied to the investigation of transformation during sample preparation as well as to study metabolism in living organism [71-73]. Kumar et al. [74] used synthesized mixtures of BTs as spike to evaluate different extraction procedures for the determination of BTs. They found that there were large differences in the extraction efficiencies of OTCs depending on the sample matrices. The study implies that for each type of sample, a certified reference material with identical matrix is required. Unfortunately such a requirement is unrealistic, calling for suitable analytical methods which are tolerable to matrix interference.

Only a few applications using species-specific isotope dilution for PhTs are at present available [75,I]. In Paper I, different extraction procedures were evaluated for the determination of PhTs (including mono-, di- and triphenyltin (MPhT, DPhT and TPhT)) in fresh-water sediment (PACS-2) using SI-SSID and conventional internal calibration. This study reveals that when employing the conventional non-ID calibration approach, the use of a certified reference material does not necessarily guarantee the reliability of analytical results for samples having the same matrix if incomplete extraction efficiency and species transformation take place. The use of multi-isotope spiking species-specific isotope dilution has been proved to be an excellent solution for the mentioned problems [76,77].

- 12 -

4.2.2 Species-specific spiking strategies

4.2.2.1 Single isotope spiking (SI) In this approach, one or more analogues enriched with the same isotope are used as spike, for

example, 124Sn-enriched MPhT, 124Sn-enriched DPhT and 124Sn-enriched TPhT are spiked into a sample for the determination of the corresponding species [I].

The SI-SSID approach could be used for routine quantification since it can, within reasonable limits, compensate for the inefficiency of the extraction (only if the spike is thoroughly mixed and equilibrated with the sample) as well as instability of instrumental response.

If species degradation occurs, the analytes and the spiked species should transform to the same extent. The degradation correction, which is based on isotopic ratioing, will be accurate only when the proportions between spiked species and those between the analytes in the sample are the same. For example, assuming that concentrations of TPhT:DPhT:MPhT (ng g-1) are 40:20:50 and the isotope-enriched spike are 80:40:100 then the isotopic ratios between the spiked and the incipient for each species are 2. If each species decomposes by 50%, concentrations of species will become 20:20:35 for the sample and 40:40:70 for the isotope-enriched spike, the isotopic ratios remain 2. When there are difference in proportions between the analyte and the spike, the correction will not work. For example, concentrations of TPhT:DPhT:MPhT (ng g-1) are 40:20:50 and the isotope-enriched spike are 80:100:10 then the isotopic ratio for each species are 2, 5 and 0.2. If each species decompose by 50%, concentrations of species are 20:20:35 for the sample and 40:70:40 for the isotope-enriched spike, the isotopic ratios become 2, 3.5 and 1.14 Correction still holds true for TPhT but fails for DPhT and MPhT.

An efficient way to tackle the problems of species transformation is to use the multi-isotope spiking approach.

4.2.2.2 Multi-isotope spiking (MI) This approach is used for two main purposes: first, to study the formation and degradation of

species occurring during sample preparation as well as in natural ecosystems or organisms and second, to correct for transformation reactions which occur during the whole analytical procedure. The spike normally contains a mixture of species, each labeled with an individual isotope. For example a mixture of 119Sn-enriched MBT, 118Sn-enriched DBT and 117Sn-enriched TBT was used for BTs determinations [78]. Tin has ten natural isotopes [79] (see in Table 4) and MI-SSID could be carried out for up to nine different species.

Table 4: Isotopic abundance (%) of natural tin

Isotope 112 114 115 116 117 118 119 120 122 124 Abundance 0.97

±0.01 0.66

±0.01 0.34

±0.01 14.54 ±0.09

7.68 ±0.07

24.22 ±0.09

8.59 ±0.04

32.58 ±0.09

4.63 ±0.03

5.79 ±0.05

Possible interconversion pathways of TPhT and its metabolite products are illustrated in Fig.

3. For environmental monitoring, only mono-, di-, and triorganotin compounds are normally taken into consideration and triple isotope spiking is employed. However, the total tin and IOT can be useful for quality assurance of speciation analysis. This has been discussed in Paper IV. The equations used for the calculation of inter-conversion factors and corrected concentrations of species using triple isotope spiking; taken phenyltin species as an example are described in the appendix.

- 13 -

Figure 3: Possible inter-conversion pathways of IOT and phenyltin species taken from Paper III

Fi is interconversion factor corresponding to the interconversion reaction i

4.2.3 Preparation of the isotope-enriched phenyltin compounds.

4.2.3.1 Synthesis of isotope enriched phenyltin compounds The application of SSID methodology in speciation analysis is at present of increasing

interest. For many applications including organotin speciation, the availability of suitable isotope enriched compounds is still scarce, which has prohibited a broader use of SSID. Sutton et al. [80] synthesized different chloride and iodide salts of isotope enriched di- and tri-butyltin, di- and triphenyltin on a small scale. Tetraorganotin compounds were produced by alkylation or arylation of tin(IV) iodide with the corresponding Grignard reagents. Di- and triorganotin as iodide or chloride salts were prepared by redistribution reactions of tin(IV) iodide and the corresponding tetraorganotin or by direct hydrolysis of tetraorganotin compounds, except for dibutyltindiiodide which was synthesized directly by reaction between tin metal and butyliodide. The synthesized OTCs were purified by precipitation as fluoride salts. One limitation of this procedure is the need for large amounts of salts hence isotope enriched tin metal to produce sufficient amounts of these compounds. Moreover, this purification procedure cannot efficiently separate different OTCs, making the multi-isotope spiking more complex. For example, TBT was found to contain a significant amount of DBT [81].

Encinar et al. [82] and Inagaki et al.[83] synthesized a mixture of BTs using direct reactions of isotope enriched tin metal and butyliodide with the presence of triethylamine as a catalyst. The synthesized BTs were then used for the determination of butyltin in environmental samples.

Kumar et al. [74] synthesized mixtures of 116Sn-enriched butyltin from 116SnI4 and butyl magnesium bromide. The authors then used the synthesized mixtures as spikes to evaluate different extraction procedures for the determination of butyltin in sediments.

In Paper I, a mixture of 124Sn-enriched PhTs was synthesized in a similar way using phenyl magnesium bromide. The yield of the synthesis was critically dependent on several factors such as the activity of Grignard reagent, the adding/mixing of the reaction solution, reaction time etc. Quenching the phenylation reaction by observing the color change of the reaction solution is very tricky although it is practical. The relative proportions of the individual phenyltin species were therefore difficult to control and changed from batch to batch. The mixture containing isotope-enriched MPhT, DPhT and TPhT can only be used for the determination of TPhT

- 14 -

employing SI-SSID. It is known that only MI SSID can correct for transformation/degradation of OTCs occurring in the sample [84].

In Paper II, three batches of isotope enriched MPhT, DPhT and TPhT were prepared using three different tin enriched isotopes. Preparative liquid chromatography was applied to isolate and purify the species (discussed in more detail in section 4.2.3.2). A modified procedure for the synthesis of isotope-enriched phenyltin as described in Paper I was used. MPhT was separated from DPhT and TPhT during the synthesis. A large volume of aqueous HBr was used to quench the phenylation reaction and to shift the mass transfer of MPhT to the aqueous phase. In this way, most of the less polar compounds, >95% DPhT and ~100% TPhT, were extracted to diethylether phase leaving ~95% MPhT in the aqueous phase.

4.2.3.2 Preparative liquid chromatography to isolate single isotope- enriched PhTs With HPLC, a sample is dissolved in a suitable solvent and injected into a column packed with

micro-particles, which are coated with a stationary phase. A mobile phase is pumped through the column at high pressure. Different components in the mixture pass through the column at different rates due to the differences in their partitioning behavior between the mobile liquid phase and the stationary phase. The compounds of interest are collected based on the retention times. The discussion in this section is focused on the use of HPLC for the “micro-scale” preparation of single PhTs. Preparative HPLC has the following advantages

i. Separation of low- or non-volatile, thermal labile compounds. ii. The stationary and mobile phase can be selected to achieve optimum separation.

Ion exchange and reversed phase HPLC are often used for separation of organotin species. In Paper II, a new procedure for separation of phenyltin species had to be developed with the goal of preserving the chemical integrity of species. In addition, the use of complexing agents had to be avoided in order to be able to prepare a spiked solution without disturbing the equilibrium of the spike and the sample.

Ion exchange chromatography is used for separation of ionisable organotin species on, for example, a silica-based cation exchange stationary phase. A suitable mobile phase with this column is a mixture of water, methanol or acetonitrile and ammonium acetate or citrate. Separation of tri- or disubstituted OTCs is quite straightforward. For monosubstituted organotin, acetic acid was used to adjust the mobile phase to pH 3 (Paper II). The separation of OTCs is not solely due to ion exchange but also involves reversed phase interaction and adsorption. TBT is affected by adsorption to silanol groups while DBT is retained via an ionic interaction to cationic functional groups and MBT is completely dependent on the adsorption to silanol active sites for retention [85]. Encinar et al. [76] successfully performed preparative separation of single enriched-isotope MBT, DBT and TPhT on this stationary phase using a mobile phase containing methanol/water/acetic acid/diammonium hydrogen citrate. The separated single BTs were found to be stable when stored in the mobile phase containing 50% AcOH for at least one year [76]. During the work of this thesis, attempts to apply similar separation conditions for preparation of single phenyltin were not successful. MPhT and TPhT overlapped, see in Fig. 4, probably because MPhT formed a single charged complex with citrate. DPhT was eluted close to the solvent front showing that there was no or very little interaction between DPhT and the stationary phase. This indicates that DPhT formed a neutral complex with, most likely, citrate ions in the mobile phase. In separate experiments, rapid degradation of DPhT and TPhT to MPhT was observed in the presence of citrate ion, see in Fig 5. The same phenomenon was also observed when DPhT was stored in the presence of tropolone. It is observed that complexing reagents destabilize PhTs and therefore their use should be avoided when dealing with PhTs.

- 15 -

Figure 4: Superimposed chromatograms of 100µL PhT compounds (100µg g-1 as Sn for each species) on the

Zorbax 300 SCX column using MeOH/water/acetic acid (70/20/10, v/v/v) containing 100mM diammonium hydrogen citrate as mobile phase and UV-VIS detector, 254nm.

Figure 5: Relative concentrations (%) of (a) DPhT and (b) TPhT and their degradation products when stored in

MeOH/water/acetic acid (70/20/10, v/v/v) containing 100mM diammonium hydrogen citrate as a function of temperature and time.

Reversed phase chromatography involves the use of a non-polar alkyl-bonded silica stationary phase and a polar mobile phase. The mobile phase is usually water, containing a suitable proportion of an organic modifier like methanol, acetonitrile or tetrahydrofuran. Ionic organotin

- 16 -

interacts with the alkyl bonded silica stationary phase at residual silanol sites. Asymmetric peaks for di- and trisubstituted compounds and total retention of monosubstituted OTCs are obtained for a C18 column and an eluent containing mixtures of water and methanol or tetrahydrofuran. Problems related to mono- and disubstituted OTCs are solved by adding tropolone and acetic acid to the eluent [54]. In this case, the interaction of highly charged OTCs with silanol sites on the silica surface is reduced due to the formation of low charged mono- and disubstituted organotin-tropolone complexes or mono- and disubstituted organotin-acetate ion-pairs. At the same time, the silanol sites of the stationary phase become less active because they are blocked by acetic acid and tropolone [44]. Successful separation a complex mixture of eight OTCs including MBT, DBT and TBT was made using a mixture of tetrahydrofuran or methanol, water, acetic acid and tropolone as mobile phases [86]. Triethylamine as ion pair reagent is used to improve the peak shape and to increase the retention time of trisubstituted OTCs, facilitating baseline separation when a mixture of butyl- and phenyltin species is analyzed [55].

Since tropolone induces the degradation of PhTs [55], its use had to be excluded. To achieve acceptable separation between all three species, MPhT was separated from DPhT and TPhT already during the synthesis (see more detail in section 4.2.3.1).

4.2.4 Species-specific isotope dilution to evaluate sample extraction procedures for PhTs

4.2.4.1 Stability of OTCs in environmental sample during storage The stability of OTC in standards and samples is affected by factors like adsorption/desorption

to the container material, evaporation, contamination and transformation. Transformation of OTCs in a sample during storage, in contrast to in standard solutions, is difficult to predict since the interaction of OTCs with the sample matrix is complicated and poorly investigated. Besides, transformation of OTCs also depends on storage conditions, the use of chemical reagents and the sample treatment procedure. Therefore, the initial sample preparation steps appear to be the “Achilles heel” of a speciation procedure in terms of preserving the speciation during the entire analytical procedure.

Several workers [87-91] found that degradation of OTCs in environmental water samples was induced by the presence of suspended matter and bacteria. Quevauviller et al. [92] recommended that analysis of the filtered water sample and the suspended particulate matter should be carried out for adequate dissolved OTC speciation in water. To prevent the microbial activity, the filtrate should be acidified with HCl or AcOH and preserved with formaldehyde [93]. Contamination and losses of the sample due to leaching, adsorption and photolysis are minimized by storing samples in glass, teflon or polycarbonate bottles in the dark at +4 oC or -20 oC [94].

No degradation or losses of BTs in air-dried and pasteurized sediment are observed after 12 months storage at temperatures between -20 oC and +20 oC. In such samples, PhT species was stable over 12 months when stored at -20 oC [95]. For butyl- and phenyltin compounds, sediment was recommended to be stored at -20 oC in the dark [96,97].

Morabito et al. [98,99] reported that BTs in mussel tissue were stable at -20 oC for 44 months while decomposition of MPhT and TPhT occurred even when stored at -20 oC. After 6 months of storage, the PhTs reached constant concentrations.

4.2.4.2 Evaluation of extraction procedures of OTCs in solid samples Speciation analysis of OTCs in solid samples using currently available techniques requires

extraction prior to measurement. Validation of extraction procedures is usually based on the

- 17 -

determination of recoveries of analytes from certified reference materials or fortified samples. Leaching/extraction of OTCs from solid matrices keeping their speciation preserved is not straightforward because the binding forces between OT and matrices are species and matrices dependent. In addition, some OT compounds are not stable during leaching/extraction, particularly if the matrix is complex. Quantitative recoveries of OTCs from such samples are therefore not always achieved. Common procedures involve leaching/extraction in low polar to polar organic solvents with occasional use of acids and complexing reagents. Tropolone and HCl, HBr or AcOH have been used to assist leaching/extraction of OTCs. Acid is used to weaken the organotin-matrices binding forces by displacing OT cations with hydrogen ions. Tropolone forms low charged complexes with mono- and disubstituted OTCs making them less polar and hence more soluble in organic solvents [54].

Abalos et al. [100] found that high concentrations of HCl (25%) and AcOH (99.7%) assisted the leaching of all BTs and PhTs but HCl caused degradation of TBT. Toluene was more efficient than n-hexane, especially for MBT and MPhT in sediment. After leaching the sample overnight in the presence of HCl (25%) or AcOH (99.7%) and tropolone, TPhT was totally decomposed.

Pellegrino et al. [101] found that MBT and MPhT were extracted at very low efficiencies by non-polar solvents. In addition, the absence of both acid and complexing reagents resulted in poor extraction efficiencies for di- and especially monosubstituted OTCs, even when strong polar organic solvents were used. The presence of acid seemed to improve the recoveries of TBT and TPhT. Polar organic solvents provided high extraction efficiency for all OTCs, probably due to a better “wettability” of the matrix.

Species transformation can be considered to be one of the most serious problems encountered during speciation analysis of phenyltin. As mentioned above, the transformation of PhTs is affected by several factors. This implies that broad variations of results might be encountered when a sample is analyzed in different laboratories at different times. This explains the lack of biological certified reference material (CRM) although attempts to make such CRMs have been made since 1998 [99]. It is of increasing important to have better understanding of the factors that induce species transformation and subsequently to establish an analytical method that can minimize the species transformation.

In Paper I, single spiking species-specific isotope dilution using a mixture of synthesized 124Sn-enriched PhTs was employed to evaluate different extraction procedures (see Table 5) for the determination of phenyltin in fresh water certified sediment BCR-646. All of the studied extraction procedures gave results in reasonable agreement with the certified values except for MPhT for which the recovery was 228% using method 5, see Fig. 2, Paper I. It is known that the use of high concentrations of HCl or HBr weakens the bonds of mono- and disubstituted organotin with the matrix and favors the formation of organotin-halide ion pairs, facilitating the leaching of these compounds. This is in line with reported observations of too high extraction efficiency (166% for MPhT) in mussel tissue [102]. In the absence of a sediment matrix, degradation of PhTs occurred in two of the methods (methods 1&5). In these two methods, diethylether containing traces of peroxides and HBr with small amount of bromine were used. The adverse effect of bromine in HBr was eliminated by the sediment matrix whereas the effect of peroxides in diethylether (see Fig 3a, Paper I) still remained. DPhT and MPhT were incompletely extracted with methods 1-4, which used low acidic leachates. However, corresponding results from the sediment BCR-646 using SI-SSID were in good agreement with the certified values. This indicates complete equilibration of the isotope-enriched spikes and natural occurring incipient analytes in the sample matrices. In method 1, degradation of TPhT did not affect the accuracy of TPhT concentration in the sample because the spiked and the

- 18 -

incipient TPhT degraded to the same extent. However, the degradation products, DPhT and MPhT changed the isotope ratios between the spiked and the incipient DPhT and MPhT. SI-SSID will provide accurate results only if the proportions of OT species in the spike and sample are the same,.

Table 5: Summarized extraction procedures used in Paper I.

Method Extracting solvent Complexing agent Acid/base 1 DEE tropolone (0.02%) - 2 MeOH-DCM - - 3 DEE tropolone (0.02%) HCl 0.2 M 4 MeOH - AcOH (75%) 5 DCM tropolone (0.04%) HBr (4.4 M)

In Paper III, the MI-SSID approach was used to study different commonly used extraction

procedures for the determination of PhTs in two reference materials, (mussel tissue BCR CRM-477 and fish tissue NIES-11). During this work, the synthesized and separated isotope enriched PhTs; 118Sn-enriched MPhT, 122Sn-enriched DPhT and 124Sn-enriched TPhT were added to the samples to investigate species transformation during sample preparation. For the evaluation of and correction for species transformation, a series of equations were developed for computing inter-conversion factors. For simplicity, transformation processes associated with inorganic tin were eliminated by assuming that PhTs were not formed from IOT. This assumption was reasonable because under the experimental conditions, even in the presence of µg g-1 amounts of IOT, no PhT was detected. An increased acidity of the extractant along with the use of tropolone resulted in degradation of DPhT and TPhT.

Harsh extraction conditions like sonication and microwave treatment not only accelerated the leaching of OTCs from solid matrices but also lead to increased degradation of DPhT and TPhT. In principle, when using MI-SSID calibration species transformations during sample preparation can be corrected for. However, correction was erroneous for harsh extraction conditions e.g. microwave exposure during extended time. Hydrogen radicals, which are believed to induce the degradation of PhTs [25,103], are formed during microwave extraction. In Paper III, using Electro Spin Resonance (ESR) and α-phenyl tert-butyl nitrone (PBN) as spin trapper, the hydrogen radicals produced during microwave extraction could be trapped and detected. Buchachenko et al. [104] observed that organotin compounds involving a chemical reaction governed by radical mechanism undergo “magnetic isotope effect” resulting in changes in isotopic composition. The erroneous results of phenyltin compounds are probably due to the isotope effect.

The combination of strong acids, complexing agents and medium to polar organic solvents seems to provide quantitative recovery of OTCs. One limitation of using polar organic solvents is the lack of selectivity hampering the derivatisation reactions due to the presence of high amounts co-extracted species. The use of low to medium polar solvents circumvents this limitation but reduces extraction efficiency.

Leaching/extraction of OTCs in alkaline medium such as tetramethylammonium hydroxide (TMAH), KOH or NaOH is also applicable to biological samples. However, the conditions for basic leaching, which are elevated temperature (60 oC) and long time (1-4 h), might give rise to phenyltin degradation. Results presented in Paper III show that DPhT in mussel tissue degraded to a large extent after leaching with TMAH.

Traditional leaching/extraction procedures, using a mechanical shaker, require several hours to be completed. Modern techniques such as ultrasound, microwave and accelerated solvent extraction have many advantages over conventional mechanical shaking techniques. Modern

- 19 -

techniques accelerate the extraction rate thus reducing the extraction time, enhancing the extraction efficiency for OTCs, especially monosubstituted organotins, and reducing the amount of solvent used. However, the severe limitation of these techniques is that harsh extraction conditions may give rise to species transformation thus destroying the speciation of organotin.

In Paper III, an extraction procedure excluding the use of complexing agents, ultrasonic agitation or microwave was recommended for speciation analysis of OTCs in solid samples. Although this procedure is time consuming compared to those employing microwaves, degradation of the PhTs was minimized (degradation factors <2%). This extraction can be used for both SI-SSID or non-SSID calibration approaches with minimal risk of poor accuracy due to species transformation.

4.3 Redistribution reactions of organotin compounds in standard solutions For most of the currently used analytical methods, the quantification of a substance of interest

is based on the comparison of the analytical signal obtained from the sample of unknown concentration with that obtained from a standard of known concentration. Consequently, a well-controlled standardization is highly critical for high accuracy and precision of analytical results.

To my knowledge, no report on the redistribution of OTCs in standard solutions during preparation and storage is available at present although redistribution of OTCs under specific conditions has been observed and applied for the production of organotin compounds [79,105, 106].

The rate of redistribution in well-defined systems depends on concentrations of OTCs, the size of substituted groups, the type of catalyst and the temperature. Ali et al. [105] and Kuivila [106] et al. used redistribution reactions to synthesize new OTCs compounds. High concentrations of OTCs, harsh reaction conditions e.g. high temperature and the presence of catalyst were important factors for high yields. Plazzogna et al. [107] studied the kinetic of redistribution reactions between R3SnMe (R=Me, Et, n-Pr, Iso-Pr and Bu) and Me2SnCl2 in methanol (MeOH), MeOH-water and propanol at 25-40 oC. They found these redistribution reactions to be of second order and governed by the size of the alkyl groups, the solvent type and the temperature. Their work implicitly shows that analysts should be cautious and certain that redistribution of OTC standards has not taken place. For preparation of OTC standard solutions, methanol is preferred since it is polar, has a low boiling point that is compatible with most of sample matrices, and easy to be vented. The use of other solvents like acetone, ethanol, toluene or hexane was also reported in a few publications [100,108-110]. Single OTC stock standards were stable at +4 oC for 6-12 months [111].

In Paper IV, the stability of single or admixed organotin standards in methanol was investigated in various storing conditions.

Single butyltin or TPhT as well as mixtures of BTs were found to be stable at +22oC during 6 months.

The redistribution of MPhT was found to be temperature and concentration dependent. Redistribution of MPhT to DPhT and IOT was observed even at -20 oC. The redistribution of MPhT was faster at higher concentrations suggesting that the reaction is following pseudo-first order kinetics. The degradation of MPhT to IOT also occurred but it was insignificant compared to the redistribution reaction.

DPhT could either degrade to MPhT or redistribute to TPhT and MPhT. These reactions could be observed by measuring the increase in the concentrations of TPhT and MPhT. The redistribution of DPhT was slow, probably because of steric hindrance for the collisions of two DPhT molecules. The degradation of DPhT was temperature dependent.

- 20 -

In a mixture containing butyl- and phenyltin compounds, besides the redistribution/degradation reactions occurring in single species solution, a new organotin species was observed to be formed. Using GC-MS detector, this compound was identified to be monobutylmonophenyltindichloride, which formed by the reaction between DPhT and MBT.

The reaction proceeded very rapidly, (particularly) at room temperature, as described following:

++++ +→+ 356

25694

2256

394 ))(()( SnHCSnHCHCSnHCSnHC

To minimize redistribution/transformation of the organotin standards as well as the time for the preparation of standards, it is recommended that

i. MPhT and DPhT must be stored separately at -20 oC or below and these compounds should be used within one month. Thawing and refreezing of such standard solutions should be avoided.

ii. When preparing standard solutions containing mono-, di- and trisubstituted butyl- and phenyltin species, single species should be added to a buffer for dilution and derivatisation should be carried out as soon as possible.

iii. For mixed BTs as well as single TPhT, there is no risk of transformation reaction and therefore these standards can be stored even at room temperature.

The results of the study of redistribution/degradation reactions of phenyltin standards gives

rise to the following question: Do synthesized single PhTs degrade during storage? If this occurs, the use of MI-SSID could be more problematic with respect to the calculation of degradation for each species. In Paper II, preparative liquid chromatography was used to produce single isotope-enriched phenyltin species and the species stability was checked for various storage intervals. A solution with high ionic strength was found to inhibit redistribution reactions (in case of MPhT) but it induced degradation reaction for TPhT and DPhT. The degradation of PhTs decreased at lower temperatures.

On the basis of the results in Paper III, it can be concluded that MPhT and DPhT should be stored at -20 oC in a solution containing water, methanol, acetic acid and sodium acetate. Such standards were stable for at least 6 months. TPhT can be stored in either buffer solution or in MeOH.

4.4 Gas chromatography atomic absorption spectrometry for OT speciation analysis

4.4.1 General AAS is a relatively inexpensive, robust and user friendly analytical technique, which is readily

available in most laboratories. It is characterized by high selectivity, and a low susceptibility to matrix interference effects. For speciation analysis, AAS is often combined with QFs for the atomization of volatile metals. QFs are inexpensive and sensitive, and they can be operated continuously, which is required for coupling with a chromatographic system. For organotin speciation analysis, GC-QFAAS has been extensively used during the last 20 years. The atomization of OTCs in a QF requires normally a hydrogen/oxygen flame. Compared to ICPMS detection techniques, QF-AAS is less sensitive but it is still used in many laboratories, especially in developing countries. By increasing pre-concentration factors for analyte species, insufficient sensitivity of QFAAS can be compensated for. As discussed above, high-resolution CSAAS has a future potential for improving the reliability and detection limits of AAS. Investigations on the capability of GC-CSAAS in comparison with conventional GC-LSAAS for speciation analysis are of interest.

- 21 -

In Paper V, the analytical performance of the line source and continuum source AAS, two systems for organotin speciation analysis, were studied and compared.

4.4.2 Optimization of atomization conditions Quartz tube properties The sensitivity of Sn determinations by QF-AAS depends, among other factors, on the

atomizers’ atomization temperature, gas phase composition, i.e. the ratio between hydrogen and air and the total gas flow rate. These are parameters which can be controlled easily by the operator. Nevertheless it is problematic to achieve long-term signal stability when using QF-AAS since tin atom formation at the relatively low QF temperatures is governed by reactions with hydrogen radicals. During successive measurements, the surface of the quartz tube is often gradually covered with sample residue, which is believed to change lifetime and density distribution of hydrogen radicals within the QF and hence atomization efficiency [112]. Comparison of the two spectrometers can therefore be biased. To minimize this problem, the experiments were always started with tubes reconditioned by HF. This treatment was repeated when sensitivity had dropped by 15 %. Conditioning was required after one- to two-hundred injections depending on the type of sample analyzed. The maximum error in the comparison caused by changes in atomization efficiency was thus less than 15%. Operating conditions for the quartz tube The density of H radicals within the quartz tube is governed by the hydrogen to air ratio, the temperature and the total gas flow rate. These parameters were chosen as factors for optimizing atomization conditions by a quadratic regression model. Table 6, shows the experimental design and values of responses corresponding to the experimental matrices. The absorbance values for TBT and TPhT, obtained with LSAAS, were selected as responses, since these species are representative also for alkyl-and alkyl-aryl substituted tin. When using absorbance as the response factor it has to be assumed that the absorbance noise is not significantly changed when using the different atomization parameters of the experimental design. This was a reasonable assumption because we found that the background emission of the atomizer, which constitutes the major experimentally dependent noise contribution, was not significantly changed for the different temperatures, hydrogen/air ratios and gas flow rates applied. The QT emission was 0.01 % and 0.03% of the xenon light emission at 286.4nm and 224.6 nm, respectively.

The “goodness of fit” values for the two responses were 0.990 and 0.946 and corresponding “goodness of prediction” values were 0.948 and 0.716. Thus, a good fit of the experimental data to the model was obtained. Optimum conditions for the two OT species were slightly different with respect to temperature and the total gas flow-rate, see Fig. 6. For TPhT, higher temperatures and gas flow-rates were needed. This is reasonable, because tabulated bond dissociation energies of Sn-C for alkyl and alkenyl are 66.1 and 83.2 kcal mol-1, respectively [113]; indicating that TPhT has a higher Sn-C bond-strength compared to TBT. Once operating within the optimum region, any minor deviation (by ±2%) of the parameters (% H2, total flow rate, atomization temperature) from the optimum conditions will not significantly change the sensitivity of OTCs, i.e. responses are still confined within the optimum contour circles. For the predicted and used optimum compromised conditions, see experimental, the loss in sensitivity for TBT and TPhT was minor, 1.9 and 2.8 percent, respectively. The optimum conditions were used with both instruments.

- 22 -

Table 6: Experimental matrices and values of responses corresponding to central composite design

Exp No H2 (%)

Total flow (mL min-1)

Temp. C0) TBT TPhT

1 -1 -1 -1 0.0095 0.0071 2 +1 -1 -1 0.0003 0 3 -1 +1 -1 0.0816 0.0479 4 +1 +1 -1 0.0172 0.0126 5 -1 -1 +1 0.0005 0 6 +1 -1 +1 0.0669 0.0376 7 -1 +1 +1 0.0774 0.0478 8 +1 +1 +1 0.0943 0.0512 9 0 0 0 0.1503 0.051 10 0 0 0 0.1525 0.053 11 0 0 0 0.1595 0.0597 12 -1.68 0 0 0.0005 0 13 + 1.68 0 0 0.0184 0 14 0 - 1.68 0 0.0542 0.0273 15 0 + 1.68 0 0.1145 0.0537 16 0 0 - 1.68 0.0006 0 17 0 0 + 1.68 0.0776 0.0584

4.4.3 Comparison between line- and continuum source AAS for organotin speciation For applications requiring background correction [110], CSAAS gave better LODs (see Table

2, Paper V) and background correction capability than LSAAS equipped with D2 background correction. The CSAAS system used could correct for background signal up to 1.8 Abs with a deviation of the corrected signal of ±0.01Abs while for LSAAS, a corresponding deviation was obtained for a background of 0.4 Abs. It has been reported that CSAAS has 2-3 orders of magnitude wider working range compared to LSAAS [114]. However, with the GC coupling this advantage could not be fully exploited due to the limited capacity of the GC column.

- 23 -

Figure 6: Contour plots for TBT and TPhT from the experimental design based on the quadratic regression model.

The numbers in the figures show the experimental peak area absorbance for 1.5 ng (as tin) of TBT and TPhT.

- 24 -

When integrating the CSAAS signal, the sensitivity, detection limit and reproducibility depended on the number of pixels used for calculation. Fig. 7 illustrates the absorption line profile of Sn at the 286.4 nm resonance line.

Figure 7: Absorption profile of Sn line at 286.4 nm, measured on HR-CSAAS

The highest sensitivity and reproducibility of the wavelength integrated absorbance should be obtained if all pixels over the absorption line profile were included for calculation, see Table 7. Table 7: Relative absorbance of OTs (491 ng g-1 as Sn for each species) and RSD, n=8, of time- and wavelength

integrated absorbance for various numbers of pixels used in CSAAS, measured at 224.6 nm and 286.4 nm. The integration time was 15 seconds for each species.

IOT TBT DBT MBT TPhT DPhT MPhT TPrT RSD

All pixels 1 1 1 1 1 1 1 1 -

CP±4 0,9431 0,9863 0,9709 0,9898 0,9561 0,9860 0,9621 0,9642 1,71

CP±3 0,9020 0,9777 0,9534 0,9870 0,9354 0,9773 0,9368 0,9503 2,95

CP±2 0,8061 0,9071 0,8808 0,9303 0,8424 0,9115 0,8539 0,8372 4,95

CP±1 0,6417 0,7139 0,6840 0,7332 0,6593 0,7126 0,6524 0,5958 6,76

224n

m

CP 0,2750 0,2810 0,2756 0,2770 0,2695 0,3059 0,2499 0,2252 8,78

All pixels 1 1 1 1 1 1 1 1 -

CP±4 0,9695 0,9719 0,9744 0,9704 0,9705 0,9712 0,9747 0,9721 0,19

CP±3 0,9587 0,9635 0,9659 0,9618 0,9588 0,9610 0,9675 0,9647 0,33

CP±2 0,8916 0,8960 0,8999 0,8944 0,8890 0,8902 0,9022 0,8962 0,52

CP±1 0,6994 0,7066 0,7093 0,7054 0,7016 0,7008 0,7132 0,7079 0,67

286n

m

CP 0,2760 0,2823 0,2835 0,2812 0,2789 0,2793 0,2838 0,2841 1,01

However, the best detection limit was achieved when only the central pixel (CP) and one

(CP±1) or two (CP±2) additional pixels on each side of the absorption profile were used (see Table 2, Paper V). The sensitivity of organotin determined by CSAAS at 224.6 nm and 286.4 nm is presented in Table 8.

- 25 -

Table 8: Characteristic masses (as pg Sn) for OT species in GC-QF-CSAAS at 224.6±0.05 nm and 286.4±0.05 nm

for various numbers of pixels used

Sensitivity (pg Sn/pm.s) TBT DBT MBT TPhT DPhT MPhT

CP 14,2 13,8 15,0 14,7 13,0 15,0

CP±1 5,8 5,7 6,1 6,0 5,3 6,2 224 nm CP±2 4,6 4,6 4,9 4,9 4,2 5,0

CP 24,6 25,3 27,7 26,1 23,8 27,5

CP±1 9,8 10,1 11,0 10,4 9,5 10,9 286 nm CP±2 7,7 7,9 8,7 8,2 7,5 8,6

For samples requiring background correction, CSAAS gives on average seven times better

LOD compared with LSAAS. In general, CSAAS provides equal or better LOD than LSAAS. In the author’s opinion, a commercially available CSAAS with multi-element capability would be very useful to allow cost-efficient simultaneous speciation analysis for species from several elements such as Sn, Hg, Pb etc. Multi-element procedures for such an approach have already been published in connection with a more expensive technique, GC-ICPMS [115].

5. Conclusions To date most of the work on organotin speciation analysis, which does not use SSID, probably

provides probably inadequate information for adequate risk assessment of ecosystems. To improve the quality of data in this field, the instability of some OTCs in standard solutions and transformation of OTCs in samples during the analytical procedure have to be better understood and mastered.

The work described in this thesis deals with the development of analytical methodology for speciation analysis of OTCs in environmental samples using both low-cost and advanced instrumentation. Conditions providing stable OTCs in standard solutions have been developed. Because some PhTs are very unstable when stored in normally used solvents, the conditions provided in this thesis are believed to be essential for obtaining reliable analytical results. In combination with the findings of others, the determination of PhTs by GC-ICPMS as presented in this work will also make it possible to study pathways of OTCs in ecosystems or in organisms. For routine work, analytical procedures for the determination of OTCs should be simple with little risk of species transformation. In addition, it should be possible to use low cost instrumentation; an aspect of importance for developing countries. To fulfil this goal, physical/chemical processes related to OTCs, which take place in the sample during the analytical procedure, need to be understood and well-controlled. In this work, MI-SSID has been used for elucidating the redistribution and transformation of OTCs. The knowledge acquired from these investigations is essential for setting up methods with minimum species transformation, which can be applied with non isotope selective detectors such as CSAAS. As demonstrated in this thesis CSAAS gives, compared to LSAAS, better detection limits with improved possibilities to accurately correct for spectral interferences.

For the most toxic OTCs, TBT and TPhT, only minor species degradation and no formation has been observed during the processing of samples. Therefore, accurate analytical results can be obtained by correcting data for incomplete recovery either by applying SSID or by non isotope selective detectors. For the latter case, sample preparation procedures should be optimized to achieve at least 70% recovery.

- 26 -

Appendix

Equations for computing inter-conversion factors of PhTs using multi-isotope spiking SSID calibration. All equations used for the computation of the concentrations of MPhT, DPhT and TPhT and the transformation factors F1 (TPhT to DPhT), F2 (DPhT to MPhT), F3 (TPhT to MPhT), F4 (DPhT to TPhT), F5 (MPhT to DPhT) and F6 (MPhT to TPhT) are given below. The following symbols are used for equations: MPhT

sN , DPhTsN and TPhT

sN are the number of moles of each species present in the sample; XspN is the number of moles of the phenyltin species X (X = MPhT, DPhT or TPhT) spiked into the

sample; isAt is the abundance of tin isotope i ( i = 118, 120, 122 or 124) in the sample; MPhT

spiAt , , DPhTspiAt ,

and TPhTspiAt , are the abundance of tin isotope i in the spiked 118Sn-enriched MPhT, 122Sn-enriched DPhT

and 124Sn-enriched TPhT, respectively. XmN ,124 , X

mN ,122 , XmN ,120 and X

mN ,118 are the number of moles of the

corresponding isotopic phenyltin species X in the blend; XmkjR ,/ is the tin isotope ratio of the reference

isotope j (j = 120) to the spiked isotope k (k = 118, 122, 124) measured by GC-ICPMS for the phenyltin species X in the blend after mass bias correction.

Basically, from the nine experimentally measured isotope ratios in the blend we can obtain nine isotope dilution equations which take into account the interconversion factors between the species: equations [5] [6]and [7] for TPhT, [11], [12]and [13] for DPhT and [17], [18]and [19] for MPhT. The mass balance equations for each isotope of each species are given below

- For isotope 124

( )( ) ( )( ) ( )( ) ]1[111 631,124,124431,124,12431,124,124,124 FFFNNFFFNNFFNNN MPhTsp

MPhTs

DPhTsp

DPhTs

TPhTsp

TPhTm

TPhTm −−++−−++−−+=

( )( ) ( )( ) ( )( ) ]2[111 542,124,124142,124,12442,124,124,124 FFFNNFFFNNFFNNN MPhTsp

MPhTs

TPhTsp

TPhTs

DPhTsp

DPhTs

DPhTm −−++−−++−−+=

( )( ) ( )( ) ( )( ) ]3[111 365,124,124265,124,12465,124,124,124 FFFNNFFFNNFFNNN TPhTsp

TPhTs

DPhTsp

DPhTs

MPhTsp

MPhTs

MPhTm −−++−−++−−+=

The mass balance equations for 122Sn-enriched phenyltin, 120Sn-enriched phenyltin and 118Sn-enriched phenyltin are similar to equations 1, 2 and 3 for 124Sn-enriched phenyltin with the subscripts “124,s” and “124,sp” are replaced by “122,s” and “122,sp”; “120,s” and “120,sp” and “118,s” and “118,sp”respectively.

TPhT

a) Isotope ratio 120/124 for TPhT

( ) ( ) ( )( ) ( ) ( ) ]4[

6,124,1244,124,124,124,124

6,120,1204,120,120,120,120

,124

,120,124/120 FNNFNNNN

FNNFNNNNNN

R MPhTsp

MPhTs

DPhTsp

DPhTs

TPhTsp

TPhTs

MPhTsp

MPhTs

DPhTsp

DPhTs

TPhTsp

TPhTs

TPhTm

TPhTmTPhT

m ++++++++++

==

Rearrange equation [4], the number of moles of incipient TPhT can be calculated by equation [5] using isotopes 120

and 124.

[ ]56,120,124,124/120

,124,124/120,1206

4,120,124,124/120

,124,124/120,1204

,120,124,124/120

,124,124/120,120

MPhTs

ssTPhT

m

MPhTsp

TPhTm

MPhTspMPhT

sp

DPhTs

ssTPhT

m

DPhTsp

TPhTm

DPhTspDPhT

spss

TPhTm

TPhTsp

TPhTm

TPhTspTPhT

spTPhTs

NFAtAtRAtRAt

NF

NFAtAtRAtRAt

NFAtAtRAtRAt

NN

−−

−+

−−

−+

−

−=

- 27 -

Similarly, the number of moles of incipient TPhT can be calculated by the equation using isotopes 120 and 122 or

isotopes 120 and 118.

b) Isotope ratio 120/122 for TPhT

[ ]66,120,122,122/120

,122,122/120,1206

4,120,122,122/120

,122,122/120,1204

,120,122,122/120

,122,122/120,120

MPhTs

ssTPhT

m

MPhTsp

TPhTm

MPhTspMPhT

sp

DPhTs

ssTPhT

m

DPhTsp

TPhTm

DPhTspDPhT

spss

TPhTm

TPhTsp

TPhTm

TPhTspTPhT

spTPhTs

NFAtAtRAtRAt

NF

NFAtAtRAtRAt

NFAtAtRAtRAt

NN

−−

−+

−−

−+

−

−=

c) Isotope ratio 120/118 for TPhT

[ ]76,120,118,118/120

,118,118/120,1206

4,120,118,118/120

,118,118/120,1204

,120,118,118/120

,118,118/120,120

MPhTs

ssTPhT

m

MPhTsp

TPhTm

MPhTspMPhT

sp

DPhTs

ssTPhT

m

DPhTsp

TPhTm

DPhTspDPhT

spss

TPhTm

TPhTsp

TPhTm

TPhTspTPhT

spTPhTs

NFAtAtRAtRAt

NF

NFAtAtRAtRAt

NFAtAtRAtRAt

NN

−−

−+

−−

−+

−

−=

d) Computation of degradation factors F4 and F6

The three isotope dilution equations for TPhT contain five unknown parameters: NsTPhT, Ns

DPhT, NsMPhT, F4

and F6. However, by combining equations [5] and [6] and equations [5] and [7] the only unknown

parameters remaining are the interconversion factors F4 and F6, which can be easily computed (two

equations with two unknowns).

• from equations [5] and [6]:

0

]8[

,120,122,122/120

,122,122/120,120

,120,124,124/120

,124,124/120,1206

,120,122,122/120

,122,122/120,120

,120,124,124/120

,124,124/120,1204

,120,122,122/120

,122,122/120,120

,120,124,124/120

,124,124/120,120

=⎟⎟⎠

⎞⎜⎜⎝

⎛

−

−−

−

−+

⎟⎟⎠

⎞⎜⎜⎝

⎛

−

−−

−

−+

⎟⎟⎠

⎞⎜⎜⎝

⎛

−

−−

−

−

ssTPhT

m

MPhTsp

TPhTm

MPhTsp

ssTPhT

m

MPhTsp

TPhTm

MPhTspMPhT

sp

ssTPhT

m

DPhTsp

TPhTm

DPhTsp

ssTPhT

m

DPhTsp

TPhTm

DPhTspDPhT

sp

ssTPhT

m

TPhTsp

TPhTm

TPhTsp

ssTPhT

m

TPhTsp

TPhTm

TPhTspTPhT

sp

AtAtRAtRAt

AtAtRAtRAt

NF

AtAtRAtRAt

AtAtRAtRAt

NF

AtAtRAtRAt

AtAtRAtRAt

N

• from equations [5] and [7]:

0

]9[

,120,118,118/120

,118,118/120,120

,120,124,124/120

,124,124/120,1206

,120,118,118/120

,118,118/120,122

,120,124,124/120

,124,124/120,1204

,120,118,118/120

,118,118/120,120

,120,124,124/120

,124,124/120,120

=⎟⎟⎠

⎞⎜⎜⎝

⎛

−

−−

−

−+

⎟⎟⎠

⎞⎜⎜⎝

⎛

−

−−

−

−+

⎟⎟⎠

⎞⎜⎜⎝

⎛

−

−−

−

−

ssTPhT

m

MPhTsp

TPhTm

MPhTsp

ssTPhT

m

MPhTsp

TPhTm

MPhTspMPhT

sp

ssTPhT

m

DPhTsp

TPhTm

DPhTsp

ssTPhT

m

DPhTsp

TPhTm

DPhTspDPhT

sp

ssTPhT

m

TPhTsp

TPhTm

TPhTsp

ssTPhT

m

TPhTsp

TPhTm

TPhTspTPhT

sp

AtAtRAtRAt

AtAtRAtRAt

NF

AtAtRAtRAt

AtAtRAtRAt

NF

AtAtRAtRAt

AtAtRAtRAt

N

- 28 -

DPhT

a) Isotope ratio 120/122 for DPhT

( ) ( ) ( )( ) ( ) ( ) ]10[

8,122,1221,122,122,122,122

8,120,1201,120,120,120,120

,122

,120,122/120 FNNFNNNN

FNNFNNNNNN

R MPhTsp

MPhTs

TPhTsp

TPhTs

DPhTsp

DPhTs

MPhTsp

MPhTs

TPhTsp

TPhTs

DPhTsp

DPhTs

DPhTm

DPhTmDPhT

m +++++

+++++==

Rearrange equation [10], the number of moles of incipient DPhT can be calculated by equation [11] using isotopes

120 and 122.

[ ]111,120,122,122/120

,122,122/120,1201

5,120,122,122/120

,122,122/120,1205

,120,122,122/120

,122,122/120,120

TPhTs

ssDPhT

m

TPhTsp

DPhTm

TPhTspTPhT

sp

MPhTs

ssDPhT

m

MPhTsp

DPhTm

MPhTspMPhT

spss

DPhTm

DPhTsp

DPhTm

DPhTspDPhT

spDPhTs

NFAtAtRAtRAt

NF

NFAtAtRAtRAt

NFAtAtRAtRAt

NN

−−

−+

−−

−+

−

−=

Similarly, the number of moles of incipient DPhT can be calculated by the equation using isotopes 120 and 124 or

isotopes 120 and 118.

b) Isotope ratio 120/124 for DPhT

[ ]121,120,124,124/120

,124,124/120,1201

5,120,124,124/120

,124,124/120,1205

,120,124,124/120

,124,124/120,120

TPhTs

ssDPhT

m

TPhTsp

DPhTm

TPhTspTPhT

sp

MPhTs

ssDPhT

m

MPhTsp

DPhTm

MPhTspMPhT

spss

DPhTm

DPhTsp

DPhTm

DPhTspDPhT

spDPhTs

NFAtAtRAtRAt

NF

NFAtAtRAtRAt

NFAtAtRAtRAt

NN

−−

−+

−−

−+

−

−=

c) Isotope ratio 120/118 for DPhT

[ ]131,120,118,118/120

,118,118/120,1201

5,120,118,118/120

,118,118/120,1205

,120,118,118/120

,118,118/120,120

TPhTs

ssDPhT

m

TPhTsp

DPhTm

TPhTspTPhT

sp

MPhTs

ssDPhT

m

MPhTsp

DPhTm

MPhTspMPhT

spss

DPhTm

DPhTsp

DPhTm

DPhTspDPhT

spDPhTs

NFAtAtRAtRAt

NF

NFAtAtRAtRAt

NFAtAtRAtRAt

NN

−−

−+

−−

−+

−

−=

d) Computation of degradation factors F1 and F5:

The interconversion factors F1 and F5 are calculated by combining equations [11] and [12] and equations

[11] and [13].

• from equations [11] and [12]:

0

]14[,

,120,124,124/120

,124,124/120,120

,120,122,122/120

,122,122/120,1201

,120,124,124/120

,124,124/120,120

,120,122122/120

,122,122/120,1205

,120,124,124/120

,124,124/120,120

,120,122,122/120

,122,122/120,120

=⎟⎟⎠

⎞⎜⎜⎝

⎛

−

−−

−

−+

⎟⎟⎠

⎞⎜⎜⎝

⎛

−

−−

−

−+

⎟⎟⎠

⎞⎜⎜⎝

⎛

−

−−

−

−

ssDPhT

m

TPhTsp

DPhTm

TPhTsp

ssDPhT

m

TPhTsp

DPhTm

TPhTspTPhT

sp

ssDPhT

m

MPhTsp

DPhTm

MPhTsp

ssDPhT

MPhTsp

DPhTm

MPhTspMPhT

sp

ssDPhT

m

DPhTsp

DPhTm

DPhTsp

ssDPhT

m

DPhTsp

DPhTm

DPhTspDPhT

sp

AtAtRAtRAt

AtAtRAtRAt

NF

AtAtRAtRAt

AtmAtRAtRAt

NF

AtAtRAtRAt

AtAtRAtRAt

N

- 29 -

• from equations [11] and [13]:

0

]15[

,120,118,118/120

,118,118/120,120

,120,122,122/120

,122,122/120,1201

,120,118,118/120

,118,118/120,120

,120,122,122/120

,122,122/120,1205

,120,118,118/120

,118,118/120,120

,120,122,122/120

,122,122/120,120

=⎟⎟⎠

⎞⎜⎜⎝

⎛

−

−−

−

−+

⎟⎟⎠

⎞⎜⎜⎝

⎛

−

−−

−

−+

⎟⎟⎠

⎞⎜⎜⎝

⎛

−

−−

−

−

ssDPhT

m

TPhTsp

DPhTm

TPhTsp

ssDPhT

m

TPhTsp

DPhTm

TPhTspTPhT

sp

ssDPhT

m

MPhTsp

DPhTm

MPhTsp

ssDPhT

m

MPhTsp

DPhTm

MPhTspMPhT

sp

ssDPhT

m

DPhTsp

DPhTm

DPhTsp

ssDPhT

m

DPhTsp

DPhTm

DPhTspDPhT

sp

AtAtRAtRAt

AtAtRAtRAt

NF

AtAtRAtRAt

AtAtRAtRAt

NF

AtAtRAtRAt

AtAtRAtRAt

N

MPhT

( ) ( ) ( )( ) ( ) ( ) ]16[

3,118,1182,118,118,118,118

3,120,1202,120,120,120,120

,118

,120,118/120 FNNFNNNN

FNNFNNNNNN

R TPhTsp

TPhTs

DPhTsp

DPhTs

MPhTsp

MPhTs

TPhTsp

TPhTs

DPhTsp

DPhTs

MPhTsp

MPhTs

MPhTm

MPhTmMPhT

m +++++

+++++==

Rearrange equation [16], the number of moles of incipient MPhT can be calculated by equation [17] using isotopes

120 and 118. a) Isotope ratio 120/118 for MPhT

[ ]172,120,118,118/120

,118,118/120,1202

3,120,118,118/120

,118,118/120,1203

,120,118,118/120

,118,118/120,120

DPhTs

ssMPhT

m

DPhTsp

MPhTm

DPhTspDPhT

sp

TPhTs

ssMPhT

m

TPhTsp

MPhTm

TPhTspTPhT

spss

MPhTm

MPhTsp

MPhTm

MPhTspMPhT

spMPhTs

NFAtAtRAtRAt

NF

NFAtAtRAtRAt

NFAtAtRAtRAt

NN

−−

−+

−−

−+

−

−=

Similarly, the number of moles of incipient MPhT can be calculated by the equation using isotopes 120 and 124 or

isotopes 120 and 122.

b) Isotope ratio 120/124 for MPhT

[ ]182,120,124,124/120

,124,124/120,1202

3,120,124,124/120

,124,124/120,1203

,120,124,124/120

,124,124/120,120

DPhTs

ssMPhT

m

DPhTsp

MPhTm

DPhTspDPhT

sp

TPhTs

ssMPhT

m

TPhTsp

MPhTm

TPhTspTPhT

spss

MPhTm

MPhTsp

MPhTm

MPhTspMPhT

spMPhTs

NFAtAtRAtRAt

NF

NFAtAtRAtRAt

NFAtAtRAtRAt

NN

−−

−+

−−

−+

−

−=

c) Isotope ratio 120/122 for MPhT

[ ]192,120,122,122/120

,122,122/120,1202

3,120,122,122/120

,122,122/120,1203

,120,122,122/120

,122,122/120,120

DPhTs

ssMPhT

m

DPhTsp

MPhTm

DPhTspDPhT

sp

TPhTs

ssMPhT

m

TPhTsp

MPhTm

TPhTspTPhT

spss

MPhTm

MPhTsp

MPhTm

MPhTspMPhT

spMPhTs

NFAtAtRAtRAt

NF

NFAtAtRAtRAt

NFAtAtRAtRAt

NN

−−

−+

−−

−+

−−

=

- 30 -

d) Computation of degradation factors F2 and F3:

The interconversion factors F2 and F3 are calculated by combining equations [17] and [18] and equations

[17] and [19].

• from equations [17] and [18]:

0

]20[

,120,124,124/120

,124,124/120,120

,120,118,118/120

,118,118/120,1202

,120,124,124/120

,124,124/120,120

,120,118,118/120

,118,118/120,1203

,120,124,124/120

,124,124/120,120

,120,118,118/120

,118,118/120,120

=⎟⎟⎠

⎞⎜⎜⎝

⎛

−

−−

−

−+

⎟⎟⎠

⎞⎜⎜⎝

⎛

−

−−

−

−+

⎟⎟⎠

⎞⎜⎜⎝

⎛

−

−−

−

−

ssMPhT

m

DPhTsp

MPhTm

DPhTsp

ssMPhT

m

DPhTsp

MPhTm

DPhTspDPhT

sp

ssMPhT

m

TPhTsp

MPhTm

TPhTsp

ssMPhT

m

TPhTsp

MPhTm

TPhTspTPhT

sp

ssMPhT

m

MPhTsp

MPhTm

MPhTsp

ssMPhT

m

MPhTsp

MPhTm

MPhTspMPhT

sp

AtAtRAtRAt

AtAtRAtRAt

NF

AtAtRAtRAt

AtAtRAtRAt

NF

AtAtRAtRAt

AtAtRAtRAt

N

• from equations [17] and [19]:

0

]21[

,120,122,122/120

,122,122/120,120

,120,118,118/120

,118,118/120,1202

,120,122,122/120

,122,122/120,120

,120,118,118/120

,118,118/120,1203

,120,122,122/120

,122,122/120,120

,120,118,118/120

,118,118/120,120

=⎟⎟⎠

⎞⎜⎜⎝

⎛

−

−−

−

−+

⎟⎟⎠

⎞⎜⎜⎝

⎛

−

−−

−

−+

⎟⎟⎠

⎞⎜⎜⎝

⎛

−

−−

−

−

ssMPhT

m

DPhTsp

MPhTm

DPhTsp

ssMPhT

m

DPhTsp

MPhTm

DPhTspDPhT

sp

ssMPhT

m

TPhTsp

MPhTm

TPhTsp

ssMPhT

m

TPhTsp

MPhTm

TPhTspTPhT

sp

ssMPhT

m

MPhTsp

MPhTm

MPhTsp

ssMPhT

m

MPhTsp

MPhTm

MPhTspMPhT

sp

AtAtRAtRAt

AtAtRAtRAt

NF

AtAtRAtRAt

AtAtRAtRAt

NF

AtAtRAtRAt

AtAtRAtRAt

N

Once the six transformation factors are calculated, the equations [5], [11] and [17] can be used for the

calculation of the three concentrations of TPhT, DPhT and MPhT as explained in the Paper III (three

equations with three unknowns).

]22[1

11

46

15

32

⎟⎟⎟

⎠

⎞

⎜⎜⎜

⎝

⎛=

⎟⎟⎟⎟

⎠

⎞

⎜⎜⎜⎜

⎝

⎛

⎟⎟⎟

⎠

⎞

⎜⎜⎜

⎝

⎛

TPhT

DPhT

MPhT

TPhTs

DPhTs

MPhTs

ZYX

N

N

N

FFFFFF

- 31 -

Acknowledgements The work presented in this thesis has been performed at the Department of Chemistry, Analytical Chemistry in Umeå University, Sweden and partly in Analytik Jena Überlingen GmbH, Germany. I would like to take the opportunity to show my appreciation to all those who have made my work possible. I wish to express my sincere gratitude to:

Professor Wolfgang Frech, my supervisor, for giving me the chance to accomplish my PhD study in his group, for enduring guidance to sharpen me in scientific thinking and writing, for all supports to make my life in Umeå better and better. It is very kind of you to offer me a piece of land in your garden and facilities to do garden work during summer time. Special thanks to Magaretta for her hospitality with very nice discussions.

Professor Anders Cedergren, my co-supervisor, for encouraging me during my study and for valuable discussions.

Dr Radziuk and all staffs at Analytik Jena for supports in CSAAS measurements in Analytik Jena GmbH, Überlingen, Germany.

Docent Prof. Micheal Sharp for linguistic check of this thesis. All old and new members of the AAS group, Johanna, Lars, Tom, Daniel, Yvon for nice

discussions and team works. Special thanks to Dr Solomon for his valuable discussions and assistance in papers II and III. Dr. Erik for mental supports whenever I lose self-confident, for enjoyable time together in his place, for sharing knowledge of toxic or eatable mushroom. Many thanks also to Salam, Gee and Ana, for their hospitality.

Lars Lundmarks for his sustainable support since 1992 when he donated us the first AAS Pye Unicam SP-1900, which drove me to atomic spectroscopic field, for valuable expression “learning by doing”, for very nice and enjoyable activities (potatoes and tomatoes planting, wood cutting, house painting, fishing, scooter running, skiing… ) in his place, Bygdsiljum. Special thanks to Gunilla, Petra, Jonas and Gunnar for their hospitality during my visits.

Special thanks to William with the Labview programming “DW acquisition”, which goes along with me in almost all my research.

The professors and senior lecturers in the Department of Analytical Chemistry; Professor Knut Irgum, Dr. Svante Åberg for the supports.

The working staffs and doctoral students in the Department of Analytical Chemistry, Svante Johnsson for his help of repairing and constructing instruments; Anita, Ann-Helene for their supports in paper work; Josefina, Petrus, Anna, Erika, Julien, Wen, Frederik, Emma and Emil for friendly environment.

Many thanks to my Vietnamese colleagues, friends and students, Duy, Duc, T. Thuy, X. Thuy, Thien&Oanh’s family, Nang&Tuyen’s family, Doanh and Loan, Son and Chung, Thu and all the Vietnamese students belonged to Lineus-Palmer exchange program during the years 2002-2006. Special thanks to Anh Mai - my teacher, my colleague – for all valuable discussions on chromatography as well as on social life.

I am grateful to my grandmothers, my parents, and brothers and sisters, uncles and aunts and their families for care and supports.

Kim Oanh, my beloved wife, for your sacrifice, endless love and care, encouragement, understanding and faithful support. You are the best!

My thesis has been sponsored through a PhD fellowship program by Vietnam Government and by Knut and Alice Wallenberg as well as Kempe Foundations, Sweden.

- 32 -

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